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To the Graduate Council: I am submitting herewith a thesis written by Audrey Matteson entitled “Quantification and Ecological Perspectives on Cyanophages and Aquatic Viruses.” I have examined the final electronic copy of this thesis for form and content and recommend that it be accepted in partial fulfillment of the requirements for the degree of Ph. D, with a major in Microbiology. Dr. Steven W. Wilelm , Major Professor We have read this dissertation and recommend its acceptance: Dr. Alison Buchan Dr. Erik Zinser Dr. Mark Radosevich Accepted for the Council: Carolyn R. Hodges Vice Provost and Dean of the Graduate School (Original signatures are on file with official student records.) Quantification and Ecological Perspectives on Cyanophages and Aquatic Viruses A Dissertation Presented for the Doctor of Philosophy Degree The University of Tennessee, Knoxville Audrey Renee Matteson May 2011 i Copyright © 2010 by Audrey Matteson. ii ACKNOWLEDGEMENTS I still cannot believe I am writing this! This whole process could not have been done on my own, and I would like to thank several people for their help throughout my years in Tennessee. First and foremost I need to thank Dr. Steven Wilhelm for being my mentor and convincing me to continue when I felt like this was the last thing I should be doing. Next, I would like to thank my committee members Dr. Alison Buchan, Dr. George Bullerjahn, Dr. Mark Radosevich, and Dr. Erik Zinser for their suggestions and assistance over the past 4 years. Many people have passed through the Wilhelm lab and have given suggestions and help over the years and although there are too many to name, they still need to be thanked. Others on the 6th floor from the Buchan and Zinser labs have also given assistance to me and I thank them as well. While these people gave me technical support, my friends and family were the ones to emotionally get me through this. I want to thank my parents, Beryl and Deb, as well as my brother, Shawn for being there when I needed them. And lastly, I want to thank my husband, Mike, for putting up with all my crying and talks of science. I know there’s no crying in science. iii ABSTRACT It must be 350 words or less and contain no special characters. iv TABLE OF CONTENTS Chapter Page Chapter 1 Literature review ............................................................................................ 1 Objectives of this Study: ............................................................................................... 17 References: .................................................................................................................... 19 Chapter 2 Estimating virus production rates in aquatic systems............................... 34 Short Abstract: .............................................................................................................. 36 Long Abstract: .............................................................................................................. 36 Protocol Text:................................................................................................................ 36 Representative Results: ................................................................................................. 39 Discussion: .................................................................................................................... 40 Acknowledgments: ....................................................................................................... 42 References: .................................................................................................................... 46 Chapter 3 Title .................................................................Error! 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Vita 1 v LIST OF TABLES Table Page Table 1.1: List of journal articles using the g20 gene marker for phylogenetic analysis. 10 Table 1.2: Estimates of virus abundances in several aquatic ecosystems. ....................... 13 Table 2.1: Table of specific reagents and equipment used. All reagents and equipment are those used by the Wilhelm and Buchan labs at the University of Tennessee. Other appropriate products may be substituted for the purpose of this work. .......... 45 vi LIST OF FIGURES Figure Page Figure 1.1: Microbial loop adapted from Fuhrman (1999). ............................................. 15 Figure 2.1: The production of virus-like particles over a 10 hour incubation at in situ conditions using epifluorescence microscopy. Samples were collected during a phytoplankton bloom off the coast of New Zealand in September of 2008. ............ 41 Figure 2.2: Schematic diagram of the work flow process for assaying virus production. The process starts with the ultrafiltration of sample water to generate virus-free water. This is completed using an ultrafiltration system. In parallel water samples are collected from the same site and the free viruses passed through a filter while the microbial community (containing a mixture of infected and non-infected cells) is retained. This community is then resuspended in the virus free water and incubated under in situ conditions. The reoccurrence rate of viruses is then monitored for the next 10 hours to determine rates of virus production. .............................................. 43 vii Chapter 1 Literature review 1 Viruses are the most abundant organisms in the world’s oceans and there are estimates of up to 4 x 1030 viruses in the oceans (Suttle 2005) which means that the coean is composed of 270 Mt of carbon just from viruses (Wilhelm and Suttle 1999). Viral ecology based on marine bacteriophages has been a relatively new field of study. It has only been in the past several decades that scientists have focused their work on the importance of bacteriophages in aquatic ecosystems. Despite slow beginnings of this field, it has quickly demonstrated their sheer abundance and role in the microbial food web and biogeochemical cycling of nutrients (Wilhelm and Matteson 2008). First Identification of Viruses In the late 1800s, the first reports of an ultramicroscopic virus named the tobacco mosaic virus, which infected the tobacco plant, was described by Dimitri Ivanowski (Ivanowski 1892; Iwanowski 1892), but his contributions were questioned since was unsure he had a new agent for infection. Martin Beijerinck was the man who received the recognition as the first scientist to describe this “soluble” agent that could be diluted and passed on to other plants (Beijerinck 1899). It was also around this time that work was done studying the cattle disease foot-andmouth disease (Loeffler and Frosch 1898). All of the works done during this time laid the foundation for virology research. To understand and believe such an agent existed at the time was very forward thinking. It is remarkable that this work was done only thirty years after Pasteur’s experiments disproving spontaneous generation (Schwartz 2001). Until the early 1900s, there had been no publications describing a viral agent capable of lysing a bacterium. In 1915, Frederick Twort published a paper which described a possible ultramicroscopic virus that infected a micrococcus species isolated from calf vaccinia lymph (Twort and Lond 1915). He believed there were pathogenic and nonpathogenic viruses, and that only pathogenic ones had been isolated previously. He used a wide variety of different media types and samples, but never came across a virus able to be grown without a host. He did, however, publish the first results on bacteriophage. In the end, he was unsure what he had come across. Since there were accounts of viruses that infected plants and animals, he admits what he had come across may be a virus which infects bacteria. He also believed it could be an enzyme produced by the bacterium. Unfortunately he did not have the funds to pursue the research, and his work was not acknowledged by the scientific community at the time. 2 The French-Canadian Felix d’Herelle was the first to call these viruses bacteriophage , or a virus “devouring bacteria,” in 1917 (d'Herelle 1917). He was able to isolate bacteriophage effective against dysentery which he obtained from people recovering from the disease (Duckworth 1976). He received much notoriety for his work (perhaps due to its proposed use as a therapeutic agent against specific diseases) unlike Twort. He also did not cite any references in his paper, leaving it to appear he was the sole discoverer of a bacteriophage. A few years later, Twort’s paper was uncovered and there was much controversy on the true discoverer of the first bacteriophage (Duckworth 1976). Discovery of Phage Infecting Marine Bacteria: The study of marine bacteriophage lagged behind those infecting human infectious diseases up to the 1950s (Wiebe and Liston 1968). It was shown that during the years of 19431950, 101 of the 157 papers on phage dealt with Escherichia coli T series of phage (Baer and Krueger 1952). Nonetheless, it was found that during the 1940s, phages isolated by Kriss and Rukina from the Black Sea were able to infect various species of terrestrial bacteria (Spencer 1959). Since no attempt was made to culture these viruses, it must be stated that perhaps these were not viral plaques but antibiotic effects or protozoan grazing (Wiebe and Liston 1968). Even more work was done during the 1950s to isolate new phage, as well as to try new methods to estimate virus production and burst size in cultured isolates. One system extensively studied was on vibriophage, phage that infect the Gamma Proteobacteria Vibrio, by Smith and Krueger (Smith and Krueger 1952, 1954). Perhaps one of the first true marine bacteriophage was documented by Spencer during the late 1950s and early 1960s (Spencer 1955, 1959, 1963). Marine bacteria were isolated from fish (often what they called luminous bacteria identical to Photobacterium phosphoreum, today known as Vibrio phosphoreum) to screen bacteriophage. He was able to isolate species of Pseudomonas, Photobacterium, and Cytophaga. He concluded that these were true marine bacteriophage in origin because they were only effective in media mimicking seawater and had lower inactivation temperatures than other terrestrial phages (Spencer 1955). Until this time, most of the work done was to try and isolate new marine bacteriophages. Their importance in the food web, its effects on community structure and as a mechanism for horizontal gene transfer were not appreciated until 1968 (Wiebe and Liston 1968). What was known about the food web 3 at the time did not include viruses or the viral shunt (Pomeroy 1974a) so the fact that viruses may have played a big part in the food web should have warranted more attention. Perhaps more attention would have been paid if it was known how viruses are abundant and ubiquitous in aquatic ecosystems. It was also thought that bacteriophage were ecologically and numerically unimportant (Wiggins and Alexander 1985). Unfortunately it was not until transmission electron microscopy (TEM) was first used to screen seawater which showed that viruses persist in higher abundances (upwards of 104 mL-1) than previously thought (Torrella and Morita 1979). In 1989, the sheer abundance of bacteriophage in the ocean was uncovered with estimates of bacteriphage up to 108 mL-1 from various aquatic sources (Bergh et al. 1989). The high abundance of bacteriophage was further confirmed and determined that many ecologically important cyanobacterial and heterotrophic bacterial species were visibly infected (approximately 5-7%) (Proctor and Fuhrman 1990; Suttle et al. 1990). They went on to claim that infection by viruses reduced primary productivity in species such as diatoms and cyanobacteria (Suttle et al. 1990). These papers showing much higher abundances sparked new interest in studying the effects of phage in terms of distribution, activity, diversity of both host (by the “Kill the Winner” method) and phage, and its impact on biogeochemical cycling in aquatic ecosystems. Methods to Enumerate Viruses: Up to this time, the most common means to enumerate viruses was by TEM analysis (Torrella and Morita 1979; Bergh et al. 1989; Proctor and Fuhrman 1990). TEM was first used to visualize viruses in 1959 (Field 1982; Goyal 1987). This method has an advantage in that morphological characteristics (such as estimates of capsid and tail size) can be determined which is often what is used to group phage into different families. TEM is also often used to determine the minimum burst sizes. Minimum burst sizes are calculated by identifying the number of virus particles found in an infected cell through TEM (Bratbak et al. 1992; Weinbauer and Peduzzi 1994) and can be used in determining the virus induced bacterial lysis in bacterial production. In terms of enumerating viruses, TEM has often proven to provide an underestimate of true abundance compared to other techniques used today (Hara et al. 1991a; Suttle 1993; Noble and Fuhrman 1998a). 4 Historically, to determine the abundance of phages able to infect a particular host scientists have used the most probable number (MPN) assay. In this method, a specific host (for cyanophages, a cyanobacterium) is propagated with dilutions of viral samples and lysis is investigated (Suttle and Chan 1993). This method allows you to study the population that is actively infectious (specific host-infecting), but contains inherent problems. Often only laboratory strains of cyanobacteria are used for the MPN assays and very crude estimates of cyanophages are determined since cyanophages often only infect a small portion of hosts (Weinbauer 2004b). Today, epifluorescence microscopy using fluorescent dyes such as SYBR Green, SYBR Gold, DAPI, or YOPRO-1 have been used more often in estimating viral abundance most likely due to its precision, quickness and cheapness (Suttle et al. 1990; Hennes and Suttle 1995b; Noble and Fuhrman 1998a; Weinbauer 2004a; Wen et al. 2004; Sandaa and Larsen 2006). All of these dyes have been used to estimate abundance, but there have been many debates as to which yield the best results. Attributes such as low background fluorescence, stability of the dye, and adequate brightness are wanted (Wommack and Colwell 2000). Like most quantitative techniques, there are pitfalls which arise from excluding large genome viruses from counts based on size, leading to an inaccurate viral enumeration (Sommaruga et al. 1995). There are also issues of counting DNA bound to colloids which would inaccurately increase the viral abundance (Kepner Jr. and Pratt 1994). Viral abundance has also been estimated by flow cytometry (Marie et al. 1999; Chen et al. 2001; Brussaard 2004). Some of the same dyes used in epifluorescence microscopy are used to sort phage from samples in this method (e.g. SYBR Green). Sample preservation and consistency of analysis have been problematic in the past, but this technique has shown promise in analyzing viral abundances in natural environments (Brussaard 2004). Some of the drawbacks of flow cytometry to enumerate viruses are similar to those for epifluorescence microscopy (Weinbauer 2004a). A review of all the strengths and weaknesses of these techniques can be found in a review by Weinbauer (2004). Quantitative PCR (qPCR) has been used to enumerate various groups of microorganisms in both terrestrial and aquatic environments. This topic has been reviewed previously (Zhang and Fang 2006). Despite the application to microorganisms, work quantifying specific groups of phage has lagged behind. Most work analyzing viral abundance in the environment using qPCR 5 has been on viruses infecting humans and viruses in wastewater treatment plants (Lamothe et al. 2003; Laverick et al. 2004; Biofill-Mas et al. 2006; Carducci et al. 2008; Meleg et al. 2008). Analyses on the ecologically important bacteriophage and cyanophage have been sparse, most likely due to high diversity. The higher the diversity, the harder it is to make specific primers for qPCR. There has been one report of analyzing cyanomyoviruses in a Norwegian coastal water station using PCR primers specific for the g20 gene (which will be described below) (Sandaa and Larsen 2006). QPCR has also been used to study cyanophages specific for the bloom-forming cyanobacterium Microcystis aeruginosa (Takashima et al. 2007; Yoshida et al. 2010). There has also been little work done with qPCR on algal viruses with only one report on the freshwater Laurentian Great Lake Ontario (Short and Short 2009). With increased sequence information available, hopefully this technique will be utilized in the near future to tease apart abundances and production rates of specific phages in various aquatic ecosystems. Cyanobacteria and Cyanophage: One of the most diverse and ecologically important groups of prokaryotes in aquatic ecosystems is the oxygenic phototrophic cyanobacteria (also called blue-green algae). They are photosynthetic prokaryotes that require carbon dioxide, light, and inorganic substrates for survival (Mur et al. 1999). At the base of the microbial food web, cyanobacteria play a vital role in the primary production in the world’s oceans and lakes (Callieri and Stockner 2002). Within the extensive research done on marine carbon fixation, it has been noted that the marine Synechococcus and Prochlorococcus species fix between 32-80% of the carbon in oligotrophic oceans (Goericke 1993; Li 1995; Liu and Campbell 1997; Rocap et al. 2002). Their contribution to primary production in freshwater ecosystems, including the Great Lakes has not been determined, but with their high abundance in these systems, it is likely to also be high. Their importance in primary production does not overshadow their evolutionary significance in molding the primordial earth into its oxygenic form we experience today. Information and research on cyanobacteria inhabiting marine environments has been abundant (Schmidt et al. 1991; Urbach et al. 1998; Partensky et al. 1999; Moore et al. 2002; Paerl 2002; Scanlan and West 2002; Zinser et al. 2006). The photosynthetic picoplankton is a major group of cyanobacteria ranging in size from 0.2 to 2 µm (Callieri, 2002). Two genera, Prochlorococcus and Synechococcus, dominate the world’s oceans, and contribute greatly to primary production 6 (Waterbury et al. 1986; Partensky et al. 1999). Although they often co-occur in the water column, their spatial distributions (Moore et al. 1998; Scanlan and West 2002), light-harvesting (Goericke and Repeta 1992; Urbach et al. 1998; Ting et al. 2002) size (Johnson and Sieburth 1979; Chisholm et al. 1988) and nitrogen assimilation strategies (López et al. 2002; Moore et al. 2002) differ. Viruses are obligate parasites and have both RNA or DNA genomes and a protein capsid coat. They are often species-specific, and the types which target cyanobacteria are known as cyanophages. The first cyanophage was discovered in 1963 which infected the filamentous cyanobacteria Plectonema, Phormidium, and Lyngbya (Safferman and Morris 1963). At the time it was believed that these cyanophages would be able to act as biological control agents against harmful cyanobacterial blooms that often persist in many freshwater ecosystems (Martin and Benson 1988; Suttle 2000). It was later shown that this is unlikely the case since many cyanobacteria have the ability to become resistant to infection and phages alone cannot inhibit the formation and duration of a bloom event (Waterbury and Valois 1993a; Suttle 2000). It was many years later that the first reports of cyanophages in a marine ecosystem were discovered in coastal waters of the Black Sea (Moisa et al. 1981; Suttle 2000). There are three different double-stranded tailed DNA cyanophage families that are in the order Caudoviridae: Myoviridae, Siphoviridae, and Podoviridae. Their morphologically different tails help characterize them into their respective families. The Myoviridae are T4-like phage and have tails that contract with a neck separating the tail from the icosahedral capsid top. The Siphoviridae are -like phage and have long non-contractile tails, while the Podoviridae are T7-like phage and have short non-contractile tails (Suttle 2000). The myoviruses and podoviruses tend to be lytic phages while siphoviruses are able to undergo lysogeny (insertion of phage DNA into the host genome) and survive as a prophage in their host (Suttle 2005). To date, more work has been done studying the diversity and abundances of viruses in marine ecosystems (Fuhrman 1999a; Wilhem and Suttle 1999) compared to freshwater. In marine waters, the most common marine viruses are the Cyanomyoviruses which are in the Myoviridae family (Hambly et al. 2001) and have also been the most commonly isolated cyanophage family. The host ranges are also different among the different families. Myoviruses tend to have the broadest host ranges and are able to infect across genera and can infect both Prochlorococcus and Synechoccus (Suttle and Chan 1993; Waterbury and Valois 1993b) while podoviruses have very narrow host ranges 7 which suggests a different host-phage relationship between the two phage families (Wang and Chen 2008). Phage Diversity: It has been proposed that marine viruses provide the largest reservoir of genetic diversity on Earth (Hambly and Suttle 2005). These studies have been inhibited by the fact that phage are only able to be propagated if a host has been cultured. Only around 0.1% of marine bacteria are able to be cultured so obviously, on a whole, very little is known about marine bacteriophage or their true diversity (Kogure et al. 1980). This, along with the fact that there is no equivalent to the 16S or 18S rRNA gene in viruses to infer evolutionary relationships or diversity (Paul et al. 2002). Other problems such as insufficient DNA concentrations, DNA restriction enzyme resistance and the inability to properly clone viral genes due to deleterious effects have hindered some molecular work (Paul et al. 2002). With the help of several whole genome sequencing efforts, several conserved genes have been identified within various families of phages and eukaryotic viruses that can help elucidate the natural diversity of unculturable phage. Currently the genomes of 558 phages have been sequenced, 21 of these being cyanophages, and have been deposited into the NCBI database. Of the 21 sequenced genomes, 17 of these were myoviruses, 3 were podoviruses, and only 1 siphovirus. One of the first polymerase chain reaction (PCR) primers developed for marine viruses was developed in 1996 (Chen and Suttle 1996a; Chen et al. 1996) and targeted the DNA polymerase gene (pol) in eukaryotic algal viruses. This quickly led to other bacteriophage-specific biomarkers to analyze phage diversity. One marker that has been widely used is the g20 gene which is a homologue of the coliphage T4 capsid portal protein (Fuller et al. 1998; Wilson et al. 1999). This gene is important in DNA packaging, viral capsid assembly and head tail junctions. The degenerate primer set CPS1 and CPS2 were developed to target the 165-bp gene specific for the Myoviridae family of cyanophage. These primers have been used for denaturing gradient gel electrophoresis (DGGE), pulse field gel electrophoresis (PFGE), clone libraries, and qPCR analysis, but the amplicon is too small for phylogenetic analysis. A second primer was developed to increase the product size to 592-bp using the same CPS1 forward primer (Zhong et al. 2002). Many studies of marine (Zhong et al. 2002; Muhling et al. 2005; Short and Suttle 2005; Sandaa and Larsen 2006), brackish (Marston and Sallee 2003; Wang and Chen 2004a), and freshwater (Dorigo et al. 8 2004; Short and Suttle 2005; Wilhelm et al. 2006b) viral communities have been investigated using both sets of these primers, and have shown much uncharacterized diversity. Despite high diversity found in these environments, similar isolates and environmental sequences have been found globally, which suggests the movement of viruses across different biomes (Chen and Suttle 1996b; Short and Suttle 2005). An exhaustive list of published articles using the g20 gene and the primer sets used can be found in Table 1.1. It has been proposed that some of the diversity exhibited may come from bacteriophage, instead of strictly cyanophages as once thought since several clades of environmental sequences have no cultured representatives. This reinforces the fact that more molecular tools need to be developed to infer true diversity of marine phage. Recently a new g20 PCR primer set has been developed and claims to target a broader range of cyanomyoviruses based on information they gained on cultured isolates (Sullivan et al. 2008). While many use the g20 gene, others have used the major capsid protein (MCP) gp23 gene to study the diversity of myoviruses, including the cyanomyoviruses. This work has shown that the T-4 like phages is a large and divergent superfamily that can be found in the world’s oceans (Filee et al. 2005; Comeau and Krisch 2008). MCP-specific primers for freshwater filamentous cyanobacteria Nostoc and Anabaena have also been made and used to analyze the diversity of freshwater myoviruses. This type of freshwater cyanomyovirus was found to be genetically very different than the phages infecting unicellular cyanobacteria (i.e. Synechococcus and Prochlorococcus) in the world’s oceans (Baker et al. 2006). Recently primers specific for the DNA polymerase gene (pol) in the Podoviridae family of cyanophages (Wang and Chen 2008) were made available. The conservation of this gene among the podoviruses demonstrated how vital it is for the replication of its genome during lytic infections. This primer set is degenerate and were constructed from sequences from nine isolated podoviruses and four environmental sequences (Wang and Chen 2008). These primers have been used across several marine regimes (both coastal and pelagic) and have shown that podoviruses, like myoviruses, are ubiquitous in the world’s oceans and are very diverse (Huang et al. 2010). Currently no PCR primers have been constructed for the Siphoviridae. 9 Table 1.1: List of journal articles using the g20 gene marker for phylogenetic analysis. g20 Gene Primer Set CPS1/CPS2 (165 bps) CPS1 GTAGWATTTTCTACATTGAYGTTGG Freshwater This Study Marine (Marston and Sallee 2003) (Fuller et al. 1998) CPS2 GGTARCCAGAAATCYTCMAGCAT (Sandaa and Larsen 2006) (Wilson et al. 2000) (Frederickson et al. 2003) (Okunishi et al. 2002) (Sullivan et al. 2008) (Zhong et al. 2002) (McDaniel et al. 2006) CPS3/CPS4 (850 bps) (Zhong et al. 2002) CPS3 TGGTAYGTYGATGGMAGA CPS4 CATWTCWTCCCAHTCTTC CPS1/CPS8 (592 bps) CPS1 (Dorigo et al. 2004) (Marston and Sallee 2003) (Wilhelm et al. 2006a) (Wang et al. 2010) (Short and Suttle 2005) (Wang and Chen 2004b) (Zhong et al. 2002) (Short and Suttle 2005) (Sullivan et al. 2008) (Desbonnet et al. 2008) GTAGWATTTTCTACATTGAYGTTGG CPS8 AAATAYTTDCCAACAWATGGA CPS1.1/CPS8.1 (549 bps) CPS1.1 (Sullivan et al. 2008) GTAGWATWTTYTAYATTGAYGTWGG CPS8.1 ARTAYTTDCCDAYRWAWGGWTC (John et al. 2010) This study 10 (Mann et al. 2003; Millard et al. 2004; Paul and Sullivan 2005; Zeidner et al. 2005; Sullivan et al. 2006; Chenard and Suttle 2008; Wilhelm and Matteson 2008). More recently it was found that this gene was not limited to just marine cyanophages, but other freshwater cyanophages infecting unicellular cyanobacteria (Chenard and Suttle 2008; Wilhelm and Matteson 2008). Two genes encoding the D1 and D2 proteins (psbA and psbD) important in Photosystem II core reaction center have been identified in many marine cyanophage isolates and from environmental samples. These genes were found after whole genome sequencing methods of bacteriophage SPM2 which infects Synechococcus species (Mann et al. 2003). This provides an interesting link of lateral gene transfer from host to virus. It has been shown that psbA is in fact expressed during cyanophage infection which is believed to increase cyanophage fitness by prolonging photosynthesis (Lindell et al. 2004; Lindell et al. 2005). These genes were shown to be cotranscribed with other vital capsid genes in the phage (Lindell et al. 2005). The prolonged photosynthesis may increase the number of phage produced, leading to higher burst sizes from increased energy production. Like the cyanophages, many PCR primers have been developed to analyze the diversity and richness of eukaryotic algal viruses. Traditionally the DNA polymerase I gene of the B family (DNA polB) has been used for Phycodnaviridae (large double-stranded DNA viruses) (Chen and Suttle 1995; Chen et al. 1996), but more recently it was suggested that the MCP gene may also be a useful biomarker for studying all eukaryotic viruses except for Emiliania huxleyi based on increased sequence information (Schroeder et al. 2002; Larsen et al. 2008). Since studying only conserved genes does not demonstrate the true diversity of a sample, viral metagenomic work has also been done to study diversity and distribution of viruses in various ecosystems. In 2002, seawater from coastal California was used to make a metagenomic library (Breitbart et al. 2002) and showed much uncharacterized diversity that included many of the double-stranded DNA viruses including algal viruses. In 2006, four geographically distinct oceanic regions were analyzed and found to have high global diversity (Angly et al. 2006). Other ecosystems such as the Chesapeake Bay (Bench et al. 2007), Yellowstone hot springs (Schoenfeld et al. 2008), Florida reclaimed water (Rosario et al. 2009) and the British Columbia (Culley et al. 2006) have been used for viral metagenomics. High diversity is often exhibited with most sequences not found in sequence databases. Although most often the results of these 11 experiments leads to unique and unidentified genes, as more work is done, others can come back to this information later and use it in their analyses. Virus Abundances As stated previously, viruses are the most abundant organism in the oceans and are pervasive in all aquatic ecosystems. The abundance of viruses across ecosystems is quite variable and often ranges from below 104 to over 108 mL-1 (Wommack and Colwell 2000) but are generally found between 105-107 mL-1. Abundances have been determined in a vast array of locations such as oceans, lakes, sediments, hot springs, and soils and a review of these abundances can be found in Weinbauer (2004). A subset of the abundances found in freshwater and marine ecosystems can be found in Table 1.2. Although abundant in both environments, viruses in freshwater systems tend to be higher, especially in more mesotrophic or eutrophic ecosystems with higher bacterial and phytoplankton biomass (Maranger and Bird 1996; DeBruyn et al. 2004; Filippini and Middelboe 2007). Virus Production Virus production rates and turnover have been analyzed in multiple environments and are important in quantifying the bacterial community that is infected with a virus. Many techniques have been used to study virus production rates, all with their own advantages and disadvantages which have led to additional problems with analyzing the datasets across the different types. Unfortunately no technique is perfect and no reliable method has been proposed to date (Suttle 2005). A few recent articles have highlighted these techniques and the issues with each (Thingstad et al. 2008; Weinbauer et al. 2010). The first method developed detected the decay rates of viruses after the addition of a chemical to stop virus production compared with untreated controls (Heldal and Bratbak 1991; Fuhrman 1999b). A second popular technique is by radiolabeling virus particles with 3H, 32P or 14C to determine the production rate of viral DNA produced over time which is similar to the method by estimating bacterial production rates. (Steward et al. 1992a; Steward et al. 1992b). The method that has become the gold standard for virus production assays is the virus reduction (or dilution) method by which free viruses are reduced and the virally infected bacterial community is concentrated and the production of viruses are analyzed microscopically over time (Wilhlem et al. 2002). 12 Table 1.2: Estimates of virus abundances in several aquatic ecosystems. Reference Lakes: (Leff et al. 1999) (DeBruyn et al. 2004) (Wilhelm et al. 2006a) (Filippini et al. 2008) (Weinbauer and Hofle 1998) (Hennes et al. 1995) (Klut and Stockner 1990) (Tapper and Hicks 1998) (Gouvea et al. 2006) (Dorigo et al. 2004) (Brum et al. 2005) (Lymer et al. 2008) (Lymer et al. 2008) (Lymer et al. 2008) (Bettarel et al. 2006) (Vanucci et al. 2005) (Nakayama et al. 2007) (Madan et al. 2005) Sampling Location Virus Abundance (mL-1) Lake Erie Lake Erie Lake Erie Lake Hallwil, Switzerland Lake Pluβsee, Germany Lake Constance, Germany Sproat Lake, BC, Canada Lake Superior Lake Ontario Lake Bourjet, France Mono Lake, CA Lake Erken, Sweden Lake Fyrsjӧn, Sweden Lake Klocktjӧn, Sweden Lake Guiers, Senegal Lake Ganzirri, Italy Japanese Paddy Field Antarctic Lakes (Säwström et al. 2007) Antarctic Lakes, Ultraoligotrophic 1 x 106-3.4 x 107 3.0 x 107- 4.1 x 108 3.0 x 106- 4.9 x 108 (surface) 1.9-9.7 x 107 (surface) 1.37 x 107 (surface) 1-4 x 107 1.5-2x 106 7.0 x 105-9.2 x 106 (surface) 1.7-7.1 x 107 5.8 x 107- 2.6 x 108 1.4 x 108-1.9 x 109 1.2-3.7 x 108 (surface) 1.8-2.9 x 108 (surface) 5.0-6.0 x 107 (surface) 8.9 x 106-1.2 x 108 5 x 104-7.5 x 108 5.6 x 106-1.2 x 109 8.9 x 106-1.2 x 108 (whole water column) 2.0 x 105- 1.6 x 106 Marine: (Parada et al. 2007) (Manini et al. 2008) (Hennes and Suttle 1995a) (Taylor et al. 2003) (Baudoux et al. 2007) (Wilhelm et al. 1998) (Wommack et al. 1992) (Rowe et al. 2008) (Wilhelm et al. 2003) (Clasen et al. 2008) (Noble and Fuhrman 1998b) (Bettarel et al. 2002) (Winter et al. 2005) (Ortmann and Suttle 2005) (Marie et al. 1999) (Marie et al. 1999) (Hara et al. 1991b) (Weinbauer and Peduzzi 1995) (Jiang and Paul 1994) (Cochlan et al. 1993) (Bratbak et al. 1996) (Bratbak et al. 1996) (Hwang and Cho 2003) Subtropical North Atlantic Ocean Mediterranean and Pacific Hydrothermal Vents Western Gulf of Mexico Cariaco Basin, Caribbean Sea North Atlantic Ocean Gulf of Mexico Chesapeake Bay, USA Sargasso Sea Southeastern Pacific Ocean Coastal Pacific Ocean Santa Monica Bay, USA Mediterranean Sea North Sea Hydrothermal Vents Equitorial Pacific Mediterranean Sea Japanese coastal and offshore waters Northern Adratic Sea Tampa Bay Coastal, USA seasonal Southern California Bight, USA Raunefjorden, Norway Osterfjorden, Norway East Sea, Korea 1.4 x 106 (100m) 1.9-8.8 x 105 (23-75m) 4.0 x 107 (offshore) 8.1 x 106-6.3 x 107 (many depths) 1.6-2.5 x 107 3.0 x 108-6.5 x 109 2.6 x 106-1.4 x 108 2.0 x 105-1.9 x 106 (surface) 2.0 x 106- 1.6 x 107 4.0 x 106-3.9 x 108 1.5 x 107 6.0 x 106-1.1 x 107 2.1 x 106-1.8 x 108 9.2 x 106-8.9 x107 5.3 x 106 2.3 x 106 1.2 x 106- 3.5 x 107 1.0 x 106- 6.0 x 107 4.8 x 106-2.0 x 107 ~1.1 x 106-1.2 x 107 (surface) 2.0-5.0 x 107 2.0-8.0 x 107 1.0 x 106- 1.0 x 107 13 Methods to concentrate the bacterial communities using dead end and tangential flow filtration (TFF) have been compared and no significant differences were found (Weinbauer et al. 2010). Very similar to this technique is a method by which virus-free water is added to a natural sample to decrease the contact rate between host and virus to determine the production rate (Evans et al. 2003). Other methods have been employed, but intrinsic problems have decreased their usage in determining the production rate in natural samples. Biogeochemical Cycling of Nutrients Viruses have shown to have an influence on bacterial communities by the regeneration of carbon and nutrients through the lysis of bacterial hosts. Approximately 10-40% of the total bacterial population is lysed daily and this process can release a high proportion of these bioavailable nutrients (Suttle and Chan 1994; Suttle 2005). The remainder of the bacterial community is grazed on by protists (Fuhrman and Noble 1995). The release of dissolved organic matter (DOM) through the lysis of hosts can be utilized by other bacteria in the system and is known as the viral shunt since it transfers carbon away from the classic grazer-driven microbial food web (Wilhelm and Suttle 1999) (Figure 1.1). An increase in bacterial production through the release of DOM from viruses has been shown previously (Gobler et al. 1997). Often these bacterial cells get consumed by grazers such as ciliates and flagellates which, in turn, get consumed by other larger grazers which moves the energy up the food chain in the process known as the ‘microbial loop’ (Pomeroy 1974b; Azam et al. 1983). Several reviews have been written to show the importance of viruses in the global carbon cycle (Fuhrman 1999b; Wilhelm and Suttle 1999; Suttle 2005). Viruses may also may a role in the remobilization of nutrients from the lysis of bacterial hosts. This remobilization may be of importance in ecosystems that are nutrient limited (most often P in freshwater systems and Fe or N in marine systems) (Falkowski 1994). The first quantitative measurement of virus-mediated release of bioavailable nutrients was from a bloom of Aureococcus anaphageffrens which showed that C, N, P, Fe and Se were released and remobilized in different quantities after the bloom termination (Gobler et al. 1997). Others have focused their work on Fe since Fe released upon viral lysis is bioavailable for other prokaryotes in the system and may support a phytoplankton community (Poorvin et al. 2004; Mioni et al. 2005). 14 Figure 1.1: Microbial loop adapted from Fuhrman (1999). 15 Overall, little is known about the viral effect on biogeochemical cycling in freshwater ecosystems. There has been one report of the release of bioavailable P from viruses in Lake Erie and showed up to 122 - 1080 nM of phosphorus was recycled per day which reveals that viruses are an important part of the biogeochemical cycling of P in the Laurentian Great Lakes (Dean et al. 2008; Wilhelm and Matteson 2008). As phosphorus is often a limiting nutrient in Lake Erie, this recycling of P provides a significant proportion of this nutrient back to the pool. Horizontal Gene Transfer Viruses are able to move and shuffle DNA through the process of conjugation, transformation and transduction (Weinbauer 2004b). They serve as reservoirs of genetic material that are important for the movement of genes through horizontal gene transfer from viruses to hosts or viruses to viruses. This shuffling of genetic material is important for evolutionary change and leads to increased diversity in both viruses and bacteria in aquatic ecosystems (Hendrix et al. 1999; Fraser et al. 2007). This increase in diversity may also lead to an increase in fitness for the host or virus. Viral metagenomic data from many different marine environments has shown a high number of bacterial-encoded genes which suggests horizontal gene transfer between viruses and their hosts (Breitbart et al. 2002; Angly et al. 2006; Dinsdale et al. 2008; Williamson et al. 2008). From whole genome sequencing of several cyanophages, it was found that a few cyanobacterial-derived genes were found in the cyanophage genome which suggests horizontal gene transfer. Comparative genome analysis showed that these genes likely were transferred from the host to the phage several times (Lindell et al. 2004). The psbA and psbD genes which encode for the D1 and D2 proteins for Photosystem II were found the genomes of several phages (Sullivan et al. 2005; Zeidner et al. 2005) and are functional during phage infection (Lindell et al. 2005). Recently a comprehensive analysis of the genomes of 16 cyanomyoviruses were compared and found a large number of genes originating from cyanobacterial hosts important in photosynthesis, carbon metabolism, phosphate stress among others (Sullivan et al. 2010). The abundance of host genes found in cyanophage genomes suggests that this phenomenon occurs often in aquatic ecosystems and may increase the fitness of phages. Other sources of DNA for gene transfer occur from the viral release of host DNA into water by a lytic event (Weinbauer 2004b). Studies have shown that 17-32% of all free dissolved 16 DNA found in aquatic viruses is from lytic events (Jiang and Paul 1995). Upon bacterial lysis, host and viral DNA are released and can be taken up by other bacteria and viruses. Work on laboratory cultures has shown that DNA released from a lytic event can be taken up by bacteriophages (Wikner et al. 1993). Viral Control of Bacteria It is well established that viruses can control the abundance and diversity of its bacterial hosts. As a particular bacterium becomes more densely populated, the contact rates between viruses and potential hosts increases which may lead to increases in lytic events in what is known as the “killing the winner” theory (Thingstad and Lignell 1997). In this, the most dominant members of a community will give rise to an increase in viruses able to infect it, allowing the less abundant (and also resistant) members to survive. This, in turn, will increase diversity in the system. Viruses are also important in the control of phytoplankton blooms. Blooms are defined as quick increases in the population of phytoplankton and are found in many different ecosystems. These blooms are often associated with toxin production which can have detrimental effects on animals and humans. 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Zhong Y, Chen F, Wilhelm SW, Poorvin L, Hodson RE (2002) Phylogenetic Diversity of Marine Cyanophage Isolates and Natural Virus Communities as Revealed by Sequences of Viral Capsid Assembly Protein Gene g20. Appl Environ Microbiol 68(4): 1576-1584. Zinser ER, Coe A, Johnson ZI, Martiny A, Fuller NJ et al. (2006) Prochlorococcus Ecotype Abundances in the North Atlantic Ocean as revealed by an Improved Quantitative PCR Method. Applied and Environmental Microbiology 72(1): 723-732. 33 Chapter 2 Estimating virus production rates in aquatic systems 34 This section is a version of a video produced in conjunction with a video journal article made through the Journal of Visualized Experiments. Matteson AR, Budinoff CR, Campbell CE, Buchan A, Wilhelm SW (2010). Estimating Virus Production Rates in Aquatic Systems. J Vis Exp. 43. http://www.jove.com/index/details.stp?id=2196, doi: 10.3791/2196 My contribution to the published video was the assistance of writing the script, setting up all equipment to be used in the video and showing a large portion of the techniques used in the video itself. My contribution to the written article produced was help with the background information, protocol, and all figures and tables made. 35 Short Abstract: The turnover rate of viruses in marine and freshwater systems can be estimated by a reduction and reoccurrence technique. The data allow researchers to infer rates of virus-mediated microbial mortality in aquatic systems. Long Abstract: Viruses are pervasive components of marine and freshwater systems, and are known to be significant agents of microbial mortality. Developing quantitative estimates of this process is critical as we can then develop better models of microbial community structure and function as well as advance our understanding of how viruses work to alter aquatic biogeochemical cycles. The virus reduction technique allows researchers to estimate the rate at which virus particles are released from the endemic microbial community. In brief, the abundance of free (extracellular) viruses is reduced in a sample while the microbial community is maintained at near ambient concentration. The microbial community is then incubated in the absence of free viruses and the rate at which viruses reoccur in the sample (through the lysis of already infected members of the community) can be quantified by epifluorescence microscopy or, in the case of specific viruses, quantitative PCR. These rates can then be used to estimate the rate of microbial mortality due to virus-mediated cell lysis. Protocol Text: 1 Ultrafiltration of seawater to generate “virus-free” water (Wilhelm and Poorvin 2001) 1.1 Approximately 20L of seawater / lakewater is collected as aseptically as possible. 1.2 Water is serially prefiltered through 142- mm diameter polycarbonate 0.8 μm filters that can be kept at -20 °C for community analysis. Larger pore size filters may be used before this step for very productive systems. 1.3 To obtain ultrafiltered water from an Amicon M12 system (Millipore) a 30 kDacutoff spiral cartridge is used to exclude all viruses, even small RNA viruses. 1.4 The samples are processed and concentrated at ~ 25% speed with ~15-16 kPa of backpressure. 36 1.5 The remaining sample of ~500 mL of water will contain a concentrated virus community (which can be saved for other experiments) while the remaining 19.5 L of virus-free water will be used for viral production assays. 1.6 After each day of use, the Amicon M12 system must be cleaned to prevent damage to the membrane of the filter cartridge. 1.7 If you are working with seawater, rinse the membrane out with at least 6L of Milli-Q water followed by a washing with 0.1N NaOH solution for 30-45 minutes. 1.8 Again rinse the cartridge with at least 6L of Milli-Q water. 1.9 When finished using the M12 system, the spiral cartridge should be stored in a 0.05M H3PO4 solution at 4°C. 2 Virus reduction method for viral production (Wilhelm et al. 2002) 2.1 Up to 500 mL of the seawater / lakewater sample with both host and viruses is obtained and placed in a sterifilter unit with a 0.2- μm nominal pore-size low protein-binding filter (e.g., Durapore)™ placed on it. 2.2 The sample is gently vacuum pressured at <200 mmHg while continually resuspending the sample using a sterile transfer pipette to inhibit bacterial cells from concentrating on the filter. 2.3 Slowly three volumes of ultrafiltrate are added to the bacterial suspension to significantly reduce the number of free viruses in the sample. 2.4 The bacterial fraction is diluted back to 500 mL with virus-free water and divided into three replicates of 150 mL each and are placed in clear 250ml polycarbonate bottles. 3 Tangential flow filtration (TFF) method for viral production(Weinbauer et al. 2002; Winget et al. 2005). (* step 3 this represents an alternative approach to step 2) 3.1 Approximately 500 mL of natural sample is collected as described above. 3.2 This sample is concentrated using a 0.2-μm nominal pore-size tangential flow filtration system. 3.3 When the bacterial fraction is reduced to approximately 10-15 mL, ultrafiltered, virus-free water is added and distributed as above. 37 3.4 The replicate bottles are incubated at in situ conditions using environmental chambers. 3.4.1 Light levels are altered to surface conditions by using bluetinted acrylic or clear acrylic with screening net to decrease light intensity. 3.4.2 Ambient surface temperatures are often obtained by using a flowing seawater deck incubator. 3.5 Samples for bacterial and viral abundance estimates are taken at time 0 with a final concentration of 2.0-2.5% sterile glutaraldehyde added into cryovials. These samples are immediately flash frozen with liquid nitrogen and stored frozen until processed. 3.5.1 If liquid nitrogen is not available, microscopy slides may be prepared and processed immediately (see procedure below) 3.3 Subsamples are collected every 2.5 hours for at least 10 hours by the method described above. 3.3.1 At this time water may be collected for quantitative PCR analysis. Up to 5 mL of the sample may be added to a cryovial with no fixative agent with immediate flash freezing in liquid nitrogen. 4 Viral Production Microscopy (Noble and Fuhrman 1998; Wen et al. 2004) 4.1 Frozen samples to be 0.02-μm filtered for microscopy should be thawed on ice. 4.2 Prepare a stock solution of SYBR Green by diluting the stock solution 1:10 with sterile water. Next, from the stock solution, prepare a working solution by adding 1mL of the stock solution to 39 mL of sterile water. A 50% glycerol, 50% phosphate buffered saline solutions (PBS, 0.05 M Na2HPO4, 0.85% NaCl, pH 7.5) and fresh 2.5% stock solution of p-phenylenediamine should also be prepared before beginning. Keep the 50% glycerol/50% PBS solution at 4°C and the p-phenylenediamine stock at -20°C in the dark until starting. Right before filtering add the p-phenylenediamine to the 50% glycerol/50% PBS to a final concentration of 0.1% to be used as the Antifade solution. 38 4.3 Place a 25 mm 0.02-μm Anodisc filter on top of a 0.45-μm MicronStep, cellulosic backing filter. Add 850 μL of the fixed sample to the top of the Anodisc and vacuum at 20 pKa until completely dried. Place 100 μL of SYBR Green working solution to a sterile Petri dish and, with the vacuum still on, carefully remove the Anodisc from the filter tower and place on the SYBR Green. Incubate the samples in the dark at room temperature for 20 minutes. Carefully remove the filter from the SYBR Green solution and wick the back of the filter with a Kimwipe to remove all residual dye. If desired, return the filter to the tower and pass up to 800 µL of 0.02-µm filtered water or sterile media through the filter to rinse off excess stain. 4.4 Add a small drop of antifade solution to a microscope slide and place a cover slip on top. Remove the cover slip and add the dried filter to the microscope slide wet with the antifade solution. Again, add a small amount of antifade solution to the cover slip and slowly place it on top of the filter, making sure to get rid of any bubbles that may form. 4.5 Immediately freeze the slides at -20° C until needed (these should be used within a few months to prevent fading and lowered virus counts) 4.6 Viruses are enumerated using fluorescence microscopy (in our case a Leica DMRXA microscope) with a wide blue filter set (Ex= 450 to 490 nm, Em= 510 nm with a suppression filter at = 510 nm). Each filter will have at least 20 fields of view counted, making sure to quantify total viruses from each field grid to ensure even distribution of viruses across the filter membrane. 4.7 Averaged rates of virus reoccurrence from the three independent replicates are then calculated and a standard deviation is determined from the production rates. Representative Results: The raw data collected by the researcher requires minimal mathematical processing to generate reoccurrence rates of virus abundance. The primary data set resulting from this study is the reoccurrence rates of virus abundance in the subsamples from the incubations. These results form independent regressions of virus abundance vs time for each of the samples. For each 39 sample the individual incubations act as one treatment, so by completing three replicateincubations the researcher can calculate rates as well as an estimate of variation (e.g., standard deviation) (see Figure 1.) One caveat of this process is that the reduction in virus abundance invariable leads to a reduction in the host cells in the sample that are carrying the virus burden. To offset this loss, enumeration of bacterial abundance from both the source (unfiltered seawater or lake water) and the T = 0 incubation sample are necessary. This information can be used to account for the percentage of cells lost: assuming that the process of reducing virus abundance is not selective for or against any members of the microbial community, this factor can then be used to estimate the in situ production rate of viruses. Discussion: A critical component in the understanding how viruses influence marine microbial communities is to determine the rate at which virus particle are produced. Given that abundances are (more or less) static in most systems (Wilhelm and Suttle 1999; Weinbauer 2004), and that viruses are removed or rendered non-infectious quickly in aquatic systems (Wilhelm et al. 1998), then production rates must be relative rapid to replace lost particles. Estimating the mortality viruses cause to the microbial community requires a knowledge of how many viruses are produced every time a virus lyses a cell (the “burst size”). Virus burst sizes in natural samples can vary greatly. Burst sizes can be determined directly by transmission electron microscopy (e.g., Weinbauer and Peduzzi 1994), but this is often beyond the capabilities of a given laboratory or not always practical. In situations where they cannot be empirically determined, literature values of 24 viruses per lytic event may be used for marine systems and 34 for freshwater systems (Parada et al. 2006). If the rate of virus production is divided by this number, the result is the abundance of cells per volume destroyed by viruses on a daily basis. The microbes lysed value can then be divided by the standing stock of bacterial abundance resulting in the virus induced mortality for the system in question: existing estimates range from few percent to nearly the entire population and are often dependent on other factors in the system in question (Wilhelm and Matteson 2008). 40 Virus-like particles (x 107)/mL-1 8 Replicate 1 Replicate 2 Replicate 3 6 4 2 0 0 2 4 6 8 10 12 Time (h) Figure 2.1: The production of virus-like particles over a 10 hour incubation at in situ conditions using epifluorescence microscopy. Samples were collected during a phytoplankton bloom off the coast of New Zealand in September of 2008. 41 To determine the percentage of total mortality this number is often multiplied by two (working from the assumption that 50% of the cells go on to reproduce and 50% of the cells are lost, Weinbauer 2004). Given that nutrient and trace element bioavailability (e.g., N, P, Fe) can limit the rate of primary productivity, and as such carbon flux, through aquatic systems, and understanding of the role of virus-driven microbial mortality in this process has become of interest to marine geochemists. Several estimates now exist that suggest viruses release a significant concentrations of nutrient elements back to the water column on a daily basis (Rowe et al. 2008; Higgins et al. 2009) and that these elements are rapidly assimilated by the microbial community (Poorvin et al. 2004; Mioni et al. 2005). The rate of nutrient flux to the environment can be determined by multiplying the number of cells destroyed by the amount of nutrient per cell (denoted “quotas”). This information can provide a critical component to our understanding of how microbial food webs function across aquatic systems. Ongoing developments: Current efforts by a series of research groups involve adapting the above strategy to enumerate specific viruses within the community and, as such, to determine how specific organisms are influenced by virus activity. To do this researchers use the quantitative polymerase chain reaction (qPCR) to estimate the abundance of specific viruses groups or families in parallel to the estimates of the total virus community. The results are then directly applied to provide estimates of virus mortality, nutrient turnover, etc for specific plankton groups. This powerful new approach will allow researchers in the coming years to dig much more deeply into processes associated with the ecology of viruses and, for the first time, to quantify the interactions of specific virus-host communities beyond the constraints of laboratory systems. Acknowledgments: The publication of this article was supported by a grant from the Office of Research at the University of Tennessee. The authors thank the previous generation of students and researchers 42 Figure 2.2: Schematic diagram of the work flow process for assaying virus production. The process starts with the ultrafiltration of sample water to generate virus-free water. This is completed using an ultrafiltration system. In parallel water samples are collected from the same site and the free viruses passed through a filter while the microbial community (containing a mixture of infected and non-infected cells) is retained. This community is then resuspended in the virus free water and incubated under in situ conditions. The reoccurrence rate of viruses is then monitored for the next 10 hours to determine rates of virus production. 43 that have worked to refine these procedures. Research was supported by grants from the National Science Foundation (NSF-0851113, NSF-0825405 and NSF-0550485). Disclosures: We have nothing to disclose. 44 Table 2.1: Table of specific reagents and equipment used. All reagents and equipment are those used by the Wilhelm and Buchan labs at the University of Tennessee. Other appropriate products may be substituted for the purpose of this work. Name of the reagent Company Catalogue number Comments (optional) 2.5% pphenylenediamine Acros 130575000 Stock for Antifade Amicon Proflux M12 system Millipore N/A Any ultrafiltration device may be used for this step SYBR Green I nucleic Invitrogen acid gel stain S-7563 Helicon S10 30kDa Filter Millipore CDUF010LT Pelicon XL filters 0.22 μm Fisher PXGVPPC50 GE 0.2 μm PCTE membrane filters (47mm) Fisher 09-732-35 Millipore Labscale Tangential Flow Filtration System Millipore XX42LSS11 0.45 μm Micronstep, Cellulosic, white plain Fisher filters (25mm) E04WP02500 GE Whatman 0.02 μm Anodisc filters Fisher (25mm) 68-09-6002 Other TFF systems may be used for this step Any epifluorescence microscope with a blue filter set may be used Leica DMRXA microscope 20L polycarbonate carboys Fisher Glycerol Fisher BP229-4 45 PBS (0.05 M Na2HPO4, 0.85% NaCl, pH 7.5) Fisher BP329-500, S640-500 50% Glutaraldehyde Fisher G151-1 Corning 2 mL Cryovials-External Thread polypropylene Fisher 09-761-71 Any cryovials may be used Corning 5 mL Cryovials-External Thread polypropylene Fisher 09-761-74 Any cryovials may be used 85% H3PO4 Fisher A242-212 Spiral Membrane storage NaOH pellets Fisher S318-3 M12 cleaning Isopore 0.8-μm poresize membrane filter (142mm) Millipore ATTP14250 Millipore Stainless Steel Pressure Filter Holder (142mm) Fisher YY30 090 00 Graduated Cylinders References: Higgins JL, Kudo I, Nishioka J, Tsuda A, Wilhelm SW (2009) The response of the virus community to a mesoscale iron fertilization in the sub-Arctic Pacific Ocean. Deep-Sea Res II 56: 2788-2795. Mioni CE, Poorvin L, Wilhelm SW (2005) Virus and siderophore-mediated transfer of available Fe between heterotrophic bacteria: characterization using a Fe-specific bioreporter. Aquat Microb Ecol 41: 233-245. Noble RT, Fuhrman JA (1998) Use of SYBR Green I for rapid epifluorescence counts of marine viruses and bacteria. Aquat Microb Ecol 14(2): 113-118. 46 Parada V, Herndl G, Weinbauer MG (2006) Viral burst size of heterotrophic prokaryotes in aquatic systems. J Mar Biol Assoc U K 86: 613-621. Poorvin L, Rinta-Kanto JM, Hutchins DA, Wilhelm SW (2004) Viral release of Fe and its bioavailability to marine plankton. Limnol Oceanogr 49(5): 1734-1741. Rowe JM, Saxton MA, Cottrell MT, DeBruyn JM, Berg GM et al. (2008) Constraints on virus production in the Sargasso Sea and North Atlantic. Aquat Microb Ecol 52: 233-244. Weinbauer MG (2004) Ecology of prokaryotic viruses. FEMS Microbiol Rev 28: 127-181. Weinbauer MG, Peduzzi P (1994) Frequency, size and distribution of bacteriophages in different marine morphotypes. Mar Ecol Prog Ser 108: 11-20. Weinbauer MG, Winter C, Hӧfle MG (2002) Reconsidering transmission electron microscopy based estimates of viral infection of bacterioplankton using conversion factors derived from natural communities. Aquatic Microbial Ecology 27: 103-110. Wen K, Ortmann AC, Suttle CA (2004) Accurate estimation of viral abundance by epifluorescence microscopy. Appl Environ Microbiol 70(7): 3862-3867. Wilhelm SW, Suttle CA (1999) Viruses and nutrient cycles in the sea. Bioscience 49(10): 781788. Wilhelm SW, Poorvin L (2001) Quantification of algal viruses in marine samples. In: Paul JH, editor. Methods in Microbiology, vol 30 Marine Microbiology: Academic Press. pp. 5366. Wilhelm SW, Matteson AR (2008) Freshwater and marine virioplankton: a brief overview of commonalities and differences. Freshw Biol 53(6): 1076-1089. Wilhelm SW, Brigden SM, Suttle CA (2002) A dilution technique for the direct measurement of viral production: a comparison in stratified and tidally mixed coastal waters. Microb Ecol 43(1): 168-173. Wilhelm SW, Weinbauer MG, Suttle CA, Jeffrey WH (1998) The role of sunlight in the removal and repair of viruses in the sea. Limnol Oceanogr 43: 586-592. Winget DM, Williamson KE, Helton RR, Wommack KE (2005) Tangential flow diafiltration: an improved technique for estimation of virioplankton production. Aquat Microb Ecol 41: 221-232. 47 VITA Audrey Renee Matteson was born in Defiance, Ohio on April 12, 1982. She graduated from high school in 2000 and began her undergraduate degree requirements at Bowling Green State University. She graduated in 2004 with a degree in Biological Sciences with a minor in Chemistry. In the fall of 2004 she entered the graduate program at BGSU in Biological Sciences with Dr. George Bullerjahn and graduated in 2006 with her Master of Science degree. She immediately began working on her doctoral degree in Microbiology at the University of Tennessee with Dr. Steven Wilhelm and she completed the requirements for the degree in December 2010. 1